viruses has been shown to display a conditional phenotype such as heat or cold sensitivity, which would facilitate investigation

Transcription

1 JOURNAL OF VIROLOGY, Feb. 1994, P Vol. 68, No X/94/$ Copyright 1994, American Society for Microbiology Clustered Charged-to-Alanine Mutagenesis of Poliovirus RNA- Dependent RNA Polymerase Yields Multiple Temperature- Sensitive Mutants Defective in RNA Synthesis SCOTT E. DIAMOND AND KARLA KIRKEGAARD* Department of Molecular, Cellular and Developmental Biology and Howard Hughes Medical Institute, University of Colorado, Boulder, Colorado Received 29 June 1993/Accepted 29 October 1993 To generate a collection of conditionally defective poliovirus mutants, clustered charged-to-alanine mutagenesis of the RNA-dependent RNA polymerase 3D was performed. Clusters of charged residues in the polymerase coding region were replaced with alanines by deoxyoligonucleotide-directed mutagenesis of a full-length poliovirus cdna clone. Following transfection of 27 mutagenized cdna clones, 10 (37%) gave rise to viruses with temperature-sensitive (ts) phenotypes. Three of the ts mutants displayed severe ts plaque reduction phenotypes, producing at least 103-fold fewer plaques at 39.5 C than at 32.5 C; the other seven mutants displayed ts small-plaque phenotypes. Constant-temperature, single-cycle infections showed defects in virus yield or RNA accumulation at the nonpermissive temperature for eight stable ts mutants. In temperature shift experiments, seven of the ts mutants showed reduced accumulation of viral RNA at the nonpermissive temperature and showed no other ts defects. The mutations responsible for the phenotypes of most of these ts mutants lie in the N-terminal third of the 3D coding region, where no well-characterized mutations responsible for viable mutants had been previously identified. Clustered charged-to-alanine mutagenesis (S. H. Bass, M. G. Mulkerrin, and J. A. Wells, Proc. Natl. Acad. Sci. USA 88: , 1991; W. F. Bennett, N. F. Paoni, B. A. Keyt, D. Botstein, J. J. S. Jones, L. Presta, F. M. Wurm, and M. J. Zoller, J. Biol. Chem. 266: , 1991; and K. F. Wertman, D. G. Drubin, and D. Botstein, Genetics 132: , 1992) is designed to target residues on the surfaces of folded proteins; thus, extragenic suppression analysis of such mutant viruses may be very useful in identifying components of the viral replication complex. RNA-dependent RNA polymerases are required by all RNA viruses, except retroviruses, for replication of their genomes. In cases for which template specificity for the viral RNA has been reconstituted in vitro, the RNA-dependent RNA polymerases have been found to function in replicase complexes (31, 77) containing other viral or host proteins. Q,3 replicase activity requires a phage-encoded protein and three proteins from its Escherichia coli host: translational elongation factors EF-Tu and EF-Ts, as well as ribosomal protein S1 (15, 74). Templatespecific negative-strand synthesis by the Saccharomyces cerevisiae double-stranded RNA virus L-A occurs only within the intact capsid (26). For brome mosaic virus, a template-specific replicase complex was found to contain a subunit of translation initiation factor eif-3 (54). Nevertheless, enzymatic function appears to reside within virally encoded proteins, some of which display nonspecific RNA-dependent RNA polymerase activity in isolation (25, 51, 59, 73) and all of which have sequence similarity in at least four regions of the primary sequence (16, 38, 53). Two of these conserved regions are also shared by DNA- and RNA-dependent DNA polymerases (23), and one of the regions of conserved sequence, containing a highly conserved YGDD motif, may be related to the YGDTD sequence conserved among DNA-dependent DNA polymerases (5). Mutagenesis of these conserved regions has usually resulted in nonviable viruses and polymerases which display greatly reduced specific activity in vitro (32, 33, 49, 58, 59, 69). None of the mutant * Corresponding author. Mailing address: Department of Molecular, Cellular and Developmental Biology, Howard Hughes Medical Institute, University of Colorado, Campus Box 347, Boulder, CO Phone: (303) Fax: (303) viruses has been shown to display a conditional phenotype such as heat or cold sensitivity, which would facilitate investigation of the altered in vivo function. Poliovirus, a positive-strand RNA virus that infects primate cells, is a member of the family Picornaviridae. Its 7,500- nucleotide (nt) single-stranded RNA genome contains one open reading frame encoding a single 247-kDa polyprotein. This polyprotein is proteolytically processed into a number of viral polypeptides (42), one of which, 3D, displays RNAdependent RNA polymerase activity in vitro (73). The 461- amino-acid sequence of 3D (42, 56) contains the consensus sequences for RNA-dependent RNA polymerases (16, 38, 53). Yet 3D polymerase, while able to elongate RNA chains, has shown no ability either to initiate RNA synthesis independently or to provide the specificity for such synthesis in vitro (22, 71). It is likely that other viral and possibly cellular proteins form, with the polymerase, a replication complex capable of both initiating RNA synthesis and achieving specificity of RNA replication. Progress has been made in identifying the viral components of such a replication complex (13, 62). The viral protein 3B (VPg) is covalently attached to the 5' ends of poliovirus RNA strands and may act as part of the priming mechanism in vivo (62). Mutational analyses have implicated the poliovirus coding regions 2B, 2C, 3A, 3B, 3C, and 3D, as well as the 3' and 5' noncoding regions, in poliovirus RNA replication: mutations at each of these loci have been shown to cause primary defects in RNA replication (1, 3, 9, 19, 28, 36, 46, 57, 61, 70). The polypeptide products of the coding regions 2B, 2C, 3A, 3B, and 3D, along with newly synthesized poliovirus RNA, were found to colocalize to the cytoplasmic surfaces of virally induced membranous vesicles (11-13). Recently, the entire 863

2 864 DIAMOND AND KIRKEGAARD J. VIROL * 27 N - I L II I I I -I--A I., --I,, *0 lui -I * I1I1U 7.5 8, *. 10- I 1-1 I i i I I Tl I U-I ItI -1w II1 13. lb. I- I ' -I i i I - F ri I F- 11 -II I I-L I Li II - 1- r 27 I 0-1 u m ~ i1- C FIG. 1. Schematic representation of the 461-amino-acid sequence of 3D, the poliovirus RNA-dependent RNA polymerase, showing the locations of charged residues and clustered charged-to-alanine mutations. The locations of acidic residues are shown by short vertical slashes below the horizontal line that represents the 461-amino-acid sequence; basic residues are shown by slashes above the line. The locations of the clustered charged-to-alanine mutations that were introduced and the plaque phenotypes of resulting mutant viruses are shown as follows: 0, mutant virus displayed wild-type phenotype; 0, no mutant virus was recovered; *, ts virus with a plaque reduction phenotype was recovered; >, ts virus with a small-plaque phenotype was recovered. Mutant AL-14 contains an additional single-nucleotide substitution, leading to the substitution of cysteine for tyrosine at amino acid 264. intracellular poliovirus replicative cycle, including virus RNAspecific positive- and negative-strand synthesis, has been shown to occur in vitro, programmed by RNA transcripts (6, 50). These in vitro reactions contain cellular membranous structures and are extremely sensitive to treatment with nonionic detergents (49a), further supporting the idea that poliovirus-specific RNA synthesis requires membrane-associated complexes. In addition to their presumed function in both positive- and negative-strand viral RNA synthesis, 3D sequences can also function as part of the fusion protein 3CD. The 3CD polypeptide is known to function as a protease, whose specificity differs from that of the 3C protease alone (14, 37, 78). In addition, 3CD protein has been shown to bind specifically, in a complex with a 36-kDa host cell protein, to RNA sequences at the 5' end of the poliovirus positive strand (3). Conditional mutants bearing mutations in the 3D coding region that display specific defects in RNA replication would be useful in the identification of 3D residues that are required for the different functions of 3D and, possibly, for the interactions between 3D and other viral and cellular proteins. Because of the high error rate of RNA-dependent RNA polymerases (64, 65, 75), single-nucleotide substitutions are often genetically unstable; more extensive genetic changes often result in nonviable viruses (17). Thus, despite considerable effort from several laboratories, only five well-defined mutations in the 3D coding region (see Fig. 7) that give rise to viable virus with measurable phenotypic defects have been reported (10, 18, 19). Mutagenesis of one residue in 3D, Asn-424, has yielded three of these mutants; viruses containing alterations in Asn-424 were found to be temperature sensitive (ts) and to encode polymerases that were not defective for RNA elongation in vitro (1, 18). A ts mutant caused by the insertion of a single leucine residue between amino acids 257 and 258 of 3D was shown, in temperature shift experiments, to display specific defects in both RNA synthesis and proteolytic processing of viral capsid proteins at the nonpermissive temperature (19). In this case, the mutant polymerase was defective for RNA elongation in vitro (19). A small-plaque poliovirus mutant, whose nonconditional mutant phenotype was caused by insertion of a single threonine residue between amino acids 354 and 355 of 3D, displayed delayed RNA synthesis (10). A mutagenesis algorithm termed clustered charged-to-alanine mutagenesis was originally used to investigate the role of surface charges in the functions of two proteins, the human growth hormone receptor (7) and S. cerevisiae cyclic AMPdependent protein kinase (29). The resulting alleles displayed interesting in vitro phenotypes: many seem to disrupt important intermolecular contacts. For example, clustered chargedto-alanine mutagenesis of the human growth hormone receptor allowed the identification of clusters of charged residues that are required for hormone binding (7). Recently, a high proportion of mutations introduced into the S. cerevisiae actin gene, ACT], using this algorithm, were shown to give rise to S. cerevisiae that was ts in vivo (76). Specifically, 44% of the mutant alleles were shown to confer ts phenotypes to yeast cells. Why should charged-residue-to-alanine mutations cause such a high proportion of mutants with ts phenotypes? Single substitutions of charged residues with alanine in other proteins have generally been shown to exert little effect on protein conformation but instead to interfere with specific in vitro activities (20, 39, 68). Most charged residues, especially those found in clusters in the primary sequence of proteins, are expected to reside on the solvent-exposed surfaces of folded proteins (20, 76) and to contribute little to overall protein stability (21). Disruption of such clusters of surface charge might not disrupt overall protein stability but instead might interfere with electrostatic or hydrogen-bonding interactions with other biomolecules or with the solvent, making these interactions more thermosensitive (2, 76). To obtain a collection of ts mutants containing mutations in the 3D coding region and to test the utility of this technique in another protein besides actin, we constructed 27 clustered charged-to-alanine mutations in the 3D RNA-dependent RNA polymerase coding region of Mahoney type 1 poliovirus. Nine of these sets of mutations conferred ts phenotypes to the mutant polioviruses; seven of these displayed specific defects in viral RNA synthesis in temperature shift experiments. These new conditional alleles define new locations in the 3D coding region and should prove useful in disrupting specific steps of RNA replication and, possibly, in disrupting specific interactions between the polymerase and other components of the replication complex.

3 VOL. 68, 1994 CHARGED-TO-ALANINE MUTAGENESIS OF POLIOVIRUS 3D 865 MATERLALS AND METHODS Cells and viruses. Wild-type Mahoney type 1 poliovirus used in these experiments was derived from a single plaque that resulted from transfection of the infectious viral cdna (55) present in a plasmid termed pmlu, a derivative of ppolio (41). The nomenclature for viral mutants suggested by Bernstein et al. (10) was employed. HeLa cells were grown either in suspension in minimal essential medium (Joklik modification) supplemented with 7% horse serum (GIBCO Laboratories) or in petri dishes in Dulbecco modified Eagle medium supplemented with 10% calf serum (GIBCO). High-titer virus stocks were prepared by infecting HeLa cells in petri dishes (Corning) at 37 C for wild-type poliovirus and at 32.5 C for mutant polioviruses. Plaque assays were performed as described previously (40). Incubations under agar at 37 and 39.5 C were continued for 48 h, and those at 32.5 C were continued for 72 h. Construction of mutant plasmids. To create subclones of 3D-encoding poliovirus sequences suitable for site-directed mutagenesis, pmlu DNA was digested with BglII, PvuII, and EcoRI (New England Biolabs), and segments of poliovirus type 1 (Mahoney) corresponding to nt 5601 to 7053, 7053 to 7512, or 5601 to 7512 were subcloned into pbluescript KS+ DNA (Stratagene) by utilizing appropriate BglII, EcoRI and PvuII sites. The new plasmids, pbsks , pbsks , and pbsks , were transformed into CJ236 (ung dut Camr), and du-containing single-strand phagemid DNA was prepared according to the commercial protocol (Stratagene) with the following modifications: phagemid particles were pelleted from the clarified supernatant only after addition of polyethylene glycol to 5% and NaCl to 0.5 M and incubation on ice for 30 min. Deoxyoligonucleotide mutagenesis was performed according to the method of Kunkel (43), except that the Klenow fragment of E. coli polymerase I (N.E.B.) was used instead of T4 polymerase. Twenty-seven clustered charged-to-alanine mutations were constructed by using the following negativesense deoxyoligonucleotides synthesized by J. Binkley (University of Colorado). In the sequences of the mutagenic deoxyoligonucleotides given below, sequences complementary to the introduced alanine codons are underlined and the nucleotides that differ from the sequence of the poliovirus negative strand are shown in boldface. Each set of mutations created a novel BstUI (CGCG) restriction endonuclease site. The mutagenic deoxyoligonucleotides used were as follows: GGACTGCTG GCGCGGCCACCCCTTCA (AL-2); CTGTCTTAAGCGCG GGTGCGTTT'lAGTG (AL-2.5); CCTCCTCAAACGCG GTCGCAAGCCTGGGA (AL-2.7); AGAAAATTGCCGCG GCAAAGGCTGTCTTAAGC (AL-2.9); AGAAAATTGC CGCGGCAAAGTCTGTC (AL-3); CTTTTCATGTACGCGG CCACTTCAGTA (AL-4); GGTCTACTGCCGCGGCCATG TACTCA (AL-5); GGCCAGCATACGCGGCTACTGCCT CT (AL-6); AGCACA'lFFlGCGCGGTGTTGATGGCTAG TGACATG (AL-6.5); CATACATGGCCGCGGCCAAGC ACATY (AL-7); AATCAAGTGCCGCGAGACCAGCAGT GCCATAC (AL-7.5); AGATGTCTCTCGCCGCGGCTC CCATTGCT (AL-8); TGTTCAAGATCGCGGCCGCCTTFC 7'FI7CCC (AL-9); TT1TCCT7AGTCGCGGCGGThI7GThI7G (AL-10); GTFTTTGCATCGCGGCAGT GTCTCTG (AL- 11); CTCCTGGGTTCGCGGCAAAAGCAGCA (AL-13); CAAACAGCTTCGCGGCCATCAATACC (AL-14); CGAA TCCGATCGCGGCAAGCACCATC (AL-15); AAGCAA TTACCGCGGCACCATAGGCA (AL-19); CCCTGAAGAA CGCGGCCAAGAATGTT (AL-22); GAAATGGGTATGC CGCGGCTGCCCTGAAG (AL-23); ATTCATGAATCGCG GCCATTGGCATT (AL-23.5); ATCTAATTGACGCGGCA ATl'lCCTTFC (AL-24); TGTTCCTAGGCGCGGCAGTCC ATCTA (AL-25); GAGAGCGAACCGCGGCCTGAGTG TTC (AL-25); ATFFI7GTTATATGCCGCGGCGCCATTG TGC (AL-27); and AGTCAAGCCACGCGGCGTACAAT GTT (AL-28). The presence of the introduced mutations was tested by digestion with BstUI. The 3D-encoding sequences of each mutagenized plasmid were excised by digestion with BglII and PvuII or with PvlII and EcoRI and cloned into the infectious poliovirus cdna clone pmlun. pmlun was constructed from pmlu by the insertion of a Notl linker into a PvuII site in the vector sequences, rendering the PvuII site at nt 7053 in the poliovirus genome unique. Screening for ts poliovirus mutants. Transfections of DNA were performed at 37 C with cationic liposomes according to the Lipofectin (GIBCO) protocol. Plates were subsequently overlaid with Dulbecco modified Eagle medium containing 1% agar prior to incubation at 32.5 C. From each DNA transfection that gave rise to viral plaques, five well-isolated plaques were isolated and screened individually by plaque assay. Phenotypes of high-titer stocks derived from these plaque isolates were confirmed by plaque assay. Sequence verification of plasmids. For each mutant cdna that gave rise to ts virus, the entire region that was derived from the pbluescript mutagenesis vectors was sequenced, using reagents and protocols obtained from a commercial kit (Sequenase; U.S. Biochemicals). Commercially synthesized deoxyoligonucleotides (Macromolecular Resources) contained the following sequences: CTGCTCTGGTTGGAAAGTTG, complementary to poliovirus positive-strand nt 5851 to 5870; TCAAACACATAGTGGAAAGC, complementary to positive-strand nt 6071 to 6090; CCCATTGCTACATAAGGGTA, complementary to positive-strand nt 6338 to 6357; TGTGAA AAGCAGCATATAGG, complementary to positive-strand nt 6565 to 6583; TTGTACAGGTGGTGTGAGTG, complementary to positive-strand nt 6794 to 6813; GTTACATTCTC CCATGTGAC, complementary to positive-strand nt 7082 to 7101; ACTCAGGATCACGTTCGCTC, identical to positivestrand nt 7214 to 7233; and CTCCGAAYTAAAGA, complementary to positive-strand nt 7427 to Sequence verification of virus. For each ts virus, the presence of the intended mutations in the mutant viral RNAs was confirmed by sequence analysis. Virus RNA purification, cdna synthesis, and PCR from the cdna were performed as described previously (35). DNA sequencing of the PCR products was performed as described above, except that primertemplate annealing was performed by freezing on dry ice. Commercially synthesized deoxyoligonucleotides (Macromolecular Resources) contained the following sequences: AGT CAAGTCGACCATGGGTGAAATCCAGTGG, the 3-terminal 15 nucleotides of which are identical to positive-strand nucleotides 5987 to 6001, and TGAGCCCGGGTTACTA AAATGAGTC, whose 3'-terminal 20 nucleotides are complementary to positive-strand nucleotides 7361 to Constant-temperature infections. HeLa cells were grown in suspension to a density of 4.0 x 105 cells per ml. For each infection, approximately 2 x 108 cells were pelleted by lowspeed centrifugation, resuspended in 2 ml of phosphatebuffered saline (PBS), and infected with wild-type or mutant virus at a multiplicity of infection (MOI) of 50 PFU per cell. After 30 min of viral adsorbtion at room temperature, cells were washed twice with PBS, resuspended in 50 ml of minimal essential medium supplemented with 10% fetal calf serum (GIBCO), split into two fractions, and incubated with stirring at 32.5 or 39.5 C. After 60 min, HEPES (N-2-hydroxyeth-

4 866 DIAMOND AND KIRKEGAARD ylpiperazine-n'-2-ethanesulfonic acid) (ph 7.5) was added to 15 mm. At the indicated times, 2-ml samples were removed, and the cells were collected by low-speed centrifugation and washed once with PBS. The cells in one half of each sample were lysed by freezing and thawing three times, cytoplasmic extracts were prepared by centrifugation at 1,600 x g for 10 min at 4 C, and plaque assays were performed at 32.5 C. The cells in the other half of each sample were collected by low-speed centrifugation, total cytoplasmic RNA was prepared, and viral positive-stranded RNA was quantified as described previously (35). The amount of radioactive negativestrand probe hybridized to each sample was determined by a radioanalytic scanner (Ambis Systems); standard curves for each set of experiments were prepared to ensure that each measurement fell within the linear range of the assay (35). Temperature shift experiments. To measure the accumulation of viral proteins, monolayers of HeLa cells in 60-mm dishes were infected with wild-type or mutant poliovirus at an MOI of 25 PFU per cell. At 7 h postinfection at 32.5 C, cells were labeled with [35S]methionine (48), and incubation was continued at either 32.5 or 39.5 C for 15 min as described previously (41). Equal amounts of radioactivity from each supernatant were analyzed by electrophoresis in sodium dodecyl sulfate-12.5% polyacrylamide gels as described previously (44). In addition, 25-ml samples of supernatants were precipitated with 10% trichloroacetic acid and filtered through nitrocellulose; the amount of precipitated radioactivity was determined by liquid scintillation counting. To analyze the processing and stability of viral proteins that were labeled at the permissive temperature, infections with wild-type and mutant viruses were performed as described above except that the 15-min labeling with [35S]methionine was performed at 32.5 C. Labeled medium was removed, the plates were washed, label-free medium containing excess unlabeled methionine was added, and incubation was continued either for 90 min at 32.5 C or for 60 min at 39.5 C. Cells were harvested, and radioactive proteins were analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS- PAGE) as described above. RNA accumulation was measured by performing infections with wild-type and mutant polioviruses at MOIs of 25 PFU per cell as described above. After 4 h of incubation at 32.5 C, half of the plates were shifted to 39.5 C and incubation was continued. Cells on individual plates were harvested at various times postshift and collected by low-speed centrifugation, total cytoplasmic RNA was prepared, and viral positive-stranded RNA was quantified by dot blot analysis as described previously (35). Bacterial expression. For bacterial expression studies, wildtype and mutant pmlun plasmids were digested with BstBI and PvuII, and the DNA fragments corresponding to nt 6010 to 7053 of the poliovirus sequence were cloned individually into ptst-3d, replacing the wild-type 3D sequences present in that plasmid. ptst-3d was a gift of T. Jarvis (Ribozyme Products, Inc.) and is similar to the bacterial expression plasmid used by Plotch et al. to express 3D polymerase in E. coli (52). The gene expression system of Studier and Moffatt was utilized to visualize plasmid-encoded proteins (66, 67). After induction with IPTG (isopropyl-p-d-thiogalactopyranoside) for 15 min at 37 C, rifampin was added to 200,ug/ml and incubation was continued at 42 C for 5 min. Cultures were then incubated at 37 C for 20 min and labeled with [35S]methionine for 5 min. Cells were washed, collected by centrifugation, and resuspended and heated in Laemmli sample buffer. Labeled proteins were displayed by SDS-PAGE as described previously (44). RNA transfection. Mutant 3D-107 and 3D-112 pmlun plasmids were digested with BglII and EcoRI, and the DNA fragments corresponding to nt 5601 to 7512 of the poliovirus sequence were cloned individually into T7-pGEM-polio, replacing the wild-type 3D sequences present in that plasmid. T7-pGEM-polio is similar to T7-polio (60, 72). Wild-type and mutant full-length infectious transcripts produced from EcoRI-linearized plasmids were introduced into HeLa monolayers by transfection (30) with cationic liposomes (GIBCO). Plates were subsequently overlaid with Dulbecco modified Eagle medium supplemented with 10% calf serum (GIBCO) and incubated either at 32.5 C for 6 h or at 39.5 C for 4 h. Plates were washed once with PBS, and monolayers were harvested into 1 ml of PBS. The cells in each sample were lysed by freezing and thawing three times, cytoplasmic extracts were prepared by centrifugation at 1,600 x g for 10 min at 4 C, and plaque assays were performed at 32.5 C. Statistical calculations. The estimate of the standard deviation of the parental population was calculated as s = X[:5(xi-Xav)2]/(n - 1). Most experiments were done in duplicate, and n = 2. In determining the ratios of two independent variables, the relative errors were added in quadrature to determine the standard deviation of the ratio zav: sz = Zav \(Sx/Xav)2+(SyYav)2, where Zav = Xav/Yav. The Student t test of the difference between two sample means was performed: t = (Xav - Yav)/\ (s12/nl)+(s22in2) with two degrees of freedom. RESULTS J. VIROL. Clustered charged-to-alanine mutagenesis of 3D coding region. The goal of these experiments was to generate a collection of ts mutants of 3D, the poliovirus RNA-dependent RNA polymerase. To this end, clusters of two or three charged residues throughout the 3D coding region were changed sequentially to alanine, much as described by Wertman et al. (76). Specifically, any sequence of five amino acids containing two or more charged residues was a candidate to have each charged residue changed to alanine. Figure 1 shows the locations of each charged residue in the 461-amino-acid coding region of 3D and of the 27 clustered charged-to-alanine mutations that were introduced. Because of the size of the 3D coding region, all possible mutations according to the algorithm of Wertman et al. (76) were not constructed. The nucleotide and amino acid changes for each clustered charged-to-alanine mutation are given in Table 1. Mutations were introduced by deoxyoligonucleotide-directed mutagenesis (43) of segments of the 3D coding region present in phagemid vectors; each clustered charged-to-alanine mutation also created a novel BstUI restriction endonuclease site to facilitate screening for introduced mutations (see Materials and Methods). Each of the clustered charged-to-alanine mutations was subsequently transferred to a full-length, otherwise wild-type poliovirus cdna clone. Mutated full-length cdnas were introduced into HeLa monolayers at 32.5 and 39.5 C by cationic liposome-mediated transfection, and the monolayers were overlaid with agar. Twelve of the mutated cdnas gave rise to plaques after incubation at 32.5 C. ts plaque phenotypes of 10 mutant viruses resulting from clustered charged-to-alanine mutagenesis. The plaque phenotypes of the 12 viable mutant viruses were determined to identify those that were ts. From each 32.5 C DNA transfection, five individual plaques were isolated, and the titers of viruses derived from each of the plaques were determined at both 32.5 and 39.5 C on HeLa monolayers. Viruses from

5 VOL. 68, 1994 CHARGED-TO-ALANINE MUTAGENESIS OF POLIOVIRUS 3D 867 TABLE 1. Clustered charged-to-alanine mutations Original Nucleotides changed Residues changed to Plaque phenotype of virus at: Designation of designation Nu0etie ch d alanine 325C 39.5AC mutant virus AL-2 A6098G, A6099C, G6100C. A6102C. A6103G K38. E39 AL-2.5 A6126C, T6127A, A6131G, G6132C D47, R49 AL-2.7 A6137G, A6138C, A6142C, A6144C, C6145G KS1, D53 AL-2.9 AL-3 AL-4 AL-5 AL-6 AL-6.5 AL-7 AL-7.5 AL-8 AL-9 AL-10 AL-11 AL-13 AL 14b AL-15 AL-19 AL-22 AL-23 AL-23.5 AL-24 AL-25 AL-26 AL-27 A6144C, A6150C, G6151C, A6153C A6150C, G6151C, A6153C A6198C, T6199C, A6201C A6209G, A6210C, A6211C, A6213C A6222C, C6224G, A6225C, C6226G A6252C, A6262C, A6264C, A6265G A6279C, G6280C, A6282C, T6283G A6300C, A6309C, A6310G A6401G, A6402C, G6403C, A6405C, A6406G C6581G, A6582C, A6584G, A6585C, A6586G A6663C, A6664C, A6666C A6747C, G6748C, A6749G, A6750C, A6751G A6969C, T6970C, A6972C, T6973G A7109G, A7110C, G7111C, A7112G, G7113C, A7114G A7128C, A7131C, A7133G, A7134C A7169G, A7170C, G7171C, A7173C, A7174G C7178G, A7179C, T7180C, A7182C, A7183G A7199G, A7200C, A7201C, A7203C, T7204G A7221C, T7222C, C7223G, A7224C, C7225G A7263C, A7264C, A7266C, A7267G, A7269C D53, E55, E56 E55, E56 D71, E72 K75, E76 D79, H80 D89, E93 E98, D99 D105, E108 Small plaques (25 to 50% of wild-type diameter) A6359G, A6360C, G6361C, A6362G, A6363C, K125, K126,,K127 A6365G, A6366C A6365G, A6366C, AA6368G, G6369C, A6370C, K127, R128,,D129- A6372C, C6373G A6392G, G6393C, A6394C, A6396C, C6397G R136, D137 K139, E140 H149, K150 E226, E227 E254, K255 D328, D329 K375, R376 AL-28 C7349G, G7350C, C7352G, G7353C, T7354G0 R455, R456 D381, E382, K383 K395, E396 H398, E399 - K405, D406 D412, H413 E426, E427, E428 a -, does not produce virus when introduced into HeLa cells by transfection. b An additional nucleotide change, A6777G, resulted in amino acid substitution Y264C. Small plaques (50 to 80% of wild-type diameter) Small plaques (25 to 50% of wild-type diameter) 1,000 times fewer plaques Small plaques (50 to 80% of wild-type diameter) Small plaques (25 to 50% of wild-type diameter) Small plaques (25 to 50% of wild-type diameter) Small plaques (25 to 50% of wild-type diameter) Small plaques (<25% of wildtype diameter) 1,000 times fewer plaques 1,000 times fewer plaques Small plaques (<25% of wildtype diameter) 3D-106 3D-107 3D-108 3D-109 3D-110 3D-111 3D-112 3D-113 3D-114 plaque isolates from the same mutant cdna behaved identically in every case. The results of these plaque assays are summarized in Fig. 1 and Table 1. Viruses obtained from transfection of AL-6.5 and AL-11 cdnas displayed wild-type plaque phenotypes and were not characterized further. The remaining 10 viable mutant viruses exhibited ts plaque phenotypes (Table 1). Seven of these ts mutant viruses formed the same number of plaques at 39.5 as at 32.5 C, but the plaques formed at the higher temperature were distinctly smaller. One of these ts small-plaque mutants, derived from AL-7.5 cdna, also formed small plaques at 32.5 C and was quite unstable phenotypically; this virus was not characterized further. The remaining three ts mutant viruses displayed severe ts plaque reduction phenotypes, forming 1,000-fold fewer plaques at 39.5 than at 32.5 C (Table 1). Thus, a high yield of ts mutants that displayed both plaque reduction and small-plaque ts phenotypes was obtained from the clustered charged-to-alanine mutagenesis. The nine stable ts mutant viruses were named (Table 1) to conform with recommended nomenclature (10): the ts plaque reduction mutants were designated 3D-107, 3D-112, and 3D- 113, and the ts small-plaque mutants were designated 3D-106, 3D-108, 3D-109, 3D-110, 3D-111, and 3D-114. High-titer stocks of each virus were prepared, and the plaque phenotypes of viruses in these stocks were compared with those of the original plaque isolates. No differences were seen; therefore, these nine ts mutant viruses could all be propagated at the permissive condition without phenotypic reversion. Figure 2

6 868 DIAMOND AND KIRKEGAARD J. VlIROL C 39.50C b Relative concentration 32.50C 1x 39.50C ix 39.50C 104x WikJ type I;, t Wid type 11, "",// 3D D-112 FIG. 2. Plaque phenotypes of wild-type poliovirus Mahoney type 1 and ts mutant viruses. (a) and ts small-plaque mutant 3D (b) and ts plaque reduction mutant 3D-112. Infected monolayers were incubated either at 32.5 C for 72 h or at 39.5 C for 48 h. shows the results of plaque assays of high-titer stocks of wild-type virus and two mutants, ts small-plaque mutant 3D- 114 (Fig. 2a) and ts plaque reduction mutant 3D-112 (Fig. 2b). Other ts small-plaque mutants displayed a range of plaque sizes at 39.5 C, with 3D-108 showing the least dramatic, although still quite apparent, ts small-plaque phenotype (Table 1). In plaque assays of the three ts plaque reduction mutants, all behaved similarly to 3D-112 (Fig. 2b), forming approximately 1,000-fold fewer plaques at 39.5 than at 32.5 C (Table 1). Those plaques that did form after the plaque-reduction mutants 3D-107, 3D-112, and 3D-113 were plated at 39.5 C were heterogeneous in size. Several such plaques were isolated and found to harbor phenotypically revertant virus, with a much lower ratio of plaque formation at 32.50C to that at 39.50C than the original ts mutant virus (data not shown). To determine whether the introduced clustered charged-toalanine mutations were responsible for the mutant phenotypes of the ts viruses, the mutagenized region of each cdna was sequenced. Depending on the location of the introduced mutations, different regions of the 3D coding region were present in the subclone that was subjected to site-directed mutagenesis. For ts viruses 3D-106 through 3D-113, the corresponding cdnas were sequenced from nt 5601 to 7053; for ts virus 3D-1 14, the corresponding cdna was sequenced from nt 7053 to the end of the poliovirus cdna. Each of the nine cdnas that gave rise to ts viruses contained the intended mutations (Table 1). One mutated cdna, 3D-113, contained an additional, unplanned nucleotide change (A to G at nt 6777) predicted to cause an amino acid change (tyrosine to cystine at amino acid 264). Thus, in addition to the clustered charged-to-alanine mutations that were introduced, another amino acid substitution may contribute to the ts phenotype of 3D-113 virus. No other changes in the mutated poliovirus cdnas were found. To determine whether the introduced clustered charged-toalanine mutations were present in the mutant viral RNAs recovered following transfection of mutant cdnas, the presence of each mutation in viral RNA was verified by sequence analysis. For each of the nine stable ts mutants in Table 2, cdna complementary to the region surrounding the introduced mutations was synthesized and subsequently amplified by PCR (35). When these PCR products, which should represent the viral RNA sequences from which the cdna was made, were sequenced, no evidence of reversion of the introduced mutations was found. Given the high error rates of RNA-dependent RNA polymerases in general (64, 65) and of the poliovirus polymerase in particular (75), the lack of additional mutations in the transfected poliovirus cdnas and the lack of reversion at the site of the mutation in the resulting viral RNAs cannot ensure that the mutant viral RNAs obtained following transfection contained no additional mutations anywhere in the genome. It is always possible that the viruses that were selected, those that formed plaques after transfection of the cdnas, arose from a minority of RNA species that contained additional mutations that increased their viability. Two lines of evidence argue against this possibility for any of the ts viruses in this study. First, the phenotypes of the five plaques selected from each cdna transfection were identical; thus, if the viral genomes in these five plaques are not identical to the transfected cdna, but instead contain mutations that allow phenotypic reversion, either identical mutations occurred and were selected multiple times or different suppressor mutations yielded viruses with identical phenotypes. In addition, the transfection efficiencies, that is, the number of plaques that arose at 32.5 C following transfection of comparable amounts of mutated and wild-type cdna-containing plasmids, were comparable for the mutated and wild-type cdnas (data not shown). Thus, if additional suppressing mutations occurred in the RNA, they were generated at a sufficiently high frequency that the number of plaques obtained from DNA transfection was not reduced. It is therefore extremely likely that the ts phenotypes of 3D-106, 3D-107, 3D-108, 3D-109, 3D-110, 3D-111, 3D-112, and 3D-114 were caused by the clustered charged-to-alanine mutations. For 3D-113, in which an additional, unintentional point mutation was found, it is not clear whether the intended set of mutations or the additional single-nucleotide substitution at nt 6777 (Table 1) was responsible for the ts phenotype. Since the 3D-1 13 virus was found to have defects in protein processing as well as in RNA synthesis (see below) and we are primarily interested in mutants with defects only in RNA synthesis, the intended and unintended mutations present in the cdna that gave rise to ts virus 3D-113 were not introduced individually into full-length poliovirus cdna clones to determine which was responsible for the ts phenotype. Reduced accumulation of virus and RNA of ts mutants

7 VOL. 68, 1994 CHARGED-TO-ALANINE MUTAGENESIS OF POLIOVIRUS 3D J E 2L ao cc z N o '- 1L Time post infection (hours) rime post infectbn (hours) FIG. 3. Single-cycle infections showing accumulation of virus and positive-strand RNA for both wild-type and 3D-112 viruses. Constanttemperature infections were performed at 32.50C, the permissive temperature (left panels), and at 39.5 C, the nonpermissive temperature (right panels), for the ts mutant virus. Virus yield was determined by plaque assay at 32.5 C, and accumulation of positive-strand RNA was measured by quantitative dot blot hybridization (35) as described in Materials and Methods. Solid lines depict wild-type virus; dashed lines depict mutant virus. Means ± standard deviations are shown. during constant-temperature infections at 39.5 C. As a first step in characterizing the ts defects of the mutants, intracellular virus production and positive-sense viral RNA accumulation of wild-type and mutant viruses were measured during constant-temperature infections at both 32.5 and 39.5 C. HeLa suspension cultures were infected with wild-type and mutant viruses at MOIs of 50 and incubated at either 32.5 or 39.5 C. Samples were taken at intervals, and cytoplasmic extracts were prepared for analysis of virus production by plaque assay and for analysis of positive-strand RNA accumulation by quantitative dot blot hybridization. Figure 3 shows the results of a representative experiment performed with wild-type virus and ts mutant 3D-112. At 32.5 C, only slight differences in virus production and RNA accumulation between the mutant and the wild type were observed. At 39.5 C, however, both virus production and RNA accumulation were reduced at least 10-fold in infections with 3D-112 compared with those in wild-type infections. Table 2 summarizes the results of these experiments for the nine stable ts mutants. Eight of the ts mutants showed temper- Virus TABLE 2. Summary of results of constant-temperature infections at permissive (32.5 C) and nonpermissive (39.5 C) temperature for ts viruses Virus yielda at: RNA accumulation' at: 32.5 C 39.5 C 32.5 C 39.5 C ts defect(s) 3D-106 wt wt 0.8 wt 0.6 wt Slightly ts for RNA accumulation 3D-107 wt 0.1 wt 0.5 wt 0.1 wt ts for virus yield and RNA accumulation 3D wt wt 0.5 wt 0.5 wt 3D-109 wt wt 0.6 wt 0.4 wt Slightly ts for RNA accumulation 3D wt 0.4 wt 1.5 wt 0.6 wt Slightly ts for virus yield and ts for RNA accumulation 3D wt 0.2 wt 0.4 wt wt ts for virus yield 3D-112 wt 0.1 wt wt 0.1 wt ts for virus yield and RNA accumulation 3D-113b wt 0.1 wt 1.3 wt 0.2 wt ts for virus yield and RNA accumulation 3D wt 0.3 wt wt 0.3 wt ts for virus yield and RNA accumulation a Maximum yield. wt, wild type. b An additional nucleotide change, A6777G, resulted in amino acid substitution Y264C.

8 870 DIAMOND AND KIRKEGAARD TABLE 3. Virus yield following RNA transfection under permissive (32.5 C) and nonpermissive (39.5 C) conditions for ts mutants 3D- 107 and 3D-112 Virus yield (PFU/ml) Virus Replicate at: Virus yield ratio (39.5-CI32.5-C) 32.5 C 39.50C 1 5, , , , , , D , , D ature-dependent reductions of various degrees in virus yield or RNA accumulation at 39.5 C, the nonpermissive temperature. The severity of the reductions in virus and RNA accumulation correlated with the severity of the ts plaque phenotypes. 3D-108, which displayed the least severe plaque phenotype, displayed no ts defect in constant-temperature infections, while the three plaque reduction mutants, 3D-107, 3D-112, and 3D-1 13, showed the most severe ts defects in constant-temperature infections. 3D-111 displayed a ts defect only in virus yield. Several possibilities could explain this: 3D-111 might have no ts defect in RNA accumulation, it might have too subtle a defect to be distinguished by this assay, or a defect might be masked by feedback from other processes during a single cycle. These possibilities were further tested by temperature shift experiments. Two ts mutants, 3D-107 and 3D-1 12, displayed an additional defect in cell entry that resulted in a delay in progression through the infectious cycle that was apparent at both temperatures in these constant-temperature, single-cycle infections (unpublished results). In the present study, however, we consider the ts phenotypes of 3D-107 and 3D-112 independently of their slow-eclipse phenotypes. This is justified because the temperature sensitivity of 3D-107 and 3D-1 12 virus production was still observed when normal cell entry steps were bypassed by initiating wild-type, 3D-107, and 3D-112 infections with RNA transcripts (Table 3). Temperature shift experiments to monitor translation, protein processing, and protein stability of ts mutant viruses. Most of the nine stable ts mutants displayed reduced accumulation of positive-strand RNA during single-cycle infections at J. VIROL C, the temperature at which they also formed smaller or fewer plaques than wild-type viruses. However, ts defects in cell entry, translation of the viral genome, processing of viral polypeptides, or stability of the viral polypeptides, by reducing the intracellular concentrations of viral proteins, could reduce the rate of RNA replication and result in an apparent RNA accumulation defect in constant-temperature experiments. Such defects might also account for the plaque phenotypes of those mutants that showed no reduction in virus or RNA accumulation in constant-temperature infections. It is also possible that defects in RNA accumulation could be masked by feedback from these processes during long periods of infection. Thus, we tested whether any of the ts mutants showed ts defects in translation, protein processing, protein stability, or RNA synthesis after temperature shift, to try to study the temperature sensitivity of each of these processes in isolation. To assess the effect of temperature shift on translation, the accumulation of labeled viral proteins after a shift from the permissive to the nonpermissive temperature was monitored. HeLa monolayers were infected with wild-type or mutant viruses and incubated for 7 h at 32.5 C, the permissive condition; at this stage of infection, translation of the majority of host cell proteins should be inhibited, and newly synthesized proteins should be almost exclusively viral (63). At 7 h postinfection, [35S]methionine was added and the incubations were either continued at 32.5 C or shifted to 39.5 C. Following 15 min of incubation, cytoplasmic extracts were prepared and the amount of trichloroacetic acid-precipitable radioactivity was determined. As shown in Table 4, none of the ts mutants exhibited a ts reduction in the accumulation of labeled viral proteins. In fact, several mutants showed significant increases in viral protein accumulation at 39.5 C after the temperature shift compared with that of the wild type. We have, at present, no explanation for this observation; it is possible that reduced rates of RNA synthesis (see below) can lead to greater utilization of viral positive strands as mrnas. SDS-PAGE analysis (Fig. 4) of all of the extracts demonstrated that, for each mutant, inhibition of host translation occurred by 7 h postinfection at 32.5 C and that the yield and distribution of viral polypeptides synthesized at 39.5 C were equivalent to those observed in wild-type infections. Thus, none of these mutants displayed a ts defect in translation. To assess the effect of temperature shift on the processing and stability of viral proteins, HeLa cell monolayers were again infected with wild-type or mutant virus and incubated for 7 h at 32.5 C. However, in these experiments, labeling with [35S]methionine was performed for 15 min at 32.5 C immediately before the temperature shifts. Medium containing excess un- Virus TABLE 4. Translation Summary of temperature shift experiments for ts mutants RNA accumulation (cpm, 39.5/32.5 C) (cpm, 39.5/32.5 C) ts defect 1.1 ± ± 0.8 3D ± ± 0.3 ts for RNA accumulation 3D ± ± 0.1 ts for RNA accumulation 3D ± ± 0.2 ts for RNA accumulation 3D ± ± 1.2 3D ± ± 0.2 ts for RNA accumulation 3D ± ± 0.3 ts for RNA accumulation 3D ± ± 0.4 ts for RNA accumulation 3D-113a 1.3 ± 0.4 Not tested 3D ± ± 0.6 ts for RNA accumulationb An additional nucleotide change, A6777G, resulted in amino acid substitution Y264C. b Different from wild type with greater than 90% confidence by Student's t test.

9 VOL. 68, 1994 CHARGED-TO-ALANINE MUTAGENESIS OF POLIOVIRUS 3D 871 3CD - 3D - VPO - vp1 - VP3-3CD - 3D - VPO - VP1 - VP3-3CD - 3D - VPO - VP1 - VP3 - Mock 3D D-109 3D-110 AB A B A B A B A B _Y,- lw Wild typo I1 A B A B A B 3D-107 3D-112 A B A B A B _. Z = i"- 40mbomp, w A 3D 3D-114 labeled methionine was then added to identical cultures, which were then incubated either for 90 min at 32.5 C or for 60 min A B at 39.5 C to chase the labeled polypeptides under permissive and nonpermissive conditions. Cytoplasmic extracts were prepared and analyzed by SDS-PAGE (Fig. 5a). Mutants 3D-107, -108, -109, -112, and -114 revealed no defects in protein processing: no novel bands or altered polypeptide mobilities were observed with or without the temperature shift. The slight differences in the extents of protein processing between infections were not temperature dependent. For four of the ts mutants, alterations in the mobilities of viral polypeptides were observed. Three mutants, 3D-106, 3D-110, and 3D-111, showed altered mobilities of all polypeptides containing the N-terminal sequences of 3D, where the mutations responsible for the mutant phenotypes mapped (Fig. 1 and Table 1). For each of these mutants, bands - coffesponding to 3D, 3CD, and 3C' displayed greater electrophoretic mobilities than their wild-type counterparts (Fig. 5a). To determine whether these mobility shifts resulted from altered electrophoretic mobility of the mutant proteins or altered sites of protein processing, the 3D coding regions from 3D-106, 3D-110, and 3D-111 were expressed in E. coli. As shown in Fig. 5b, the observed mobilities of mutant and wild-type 3D were identical in virus-infected and bacterial cells. Furthermore, sequence analysis excluded the possibility that small deletions had been introduced into the 3D coding sequences. Therefore, the mobility shifts of 3D N-terminuscontaining polypeptides of mutants 3D-106, 3D-110, and 3D- 111 were not due to defects in protein processing but are intrinsic to the mutant polypeptides, possibly reflecting altered extents of SDS binding. Another ts mutant, 3D-113, displayed a novel band below that of capsid polypeptide VP1 at both temperatures (Fig. 5a). Although the origin of this band has not yet been determined, it is clear that 3D-113 mutant viruses, unlike the other ts mutants, displayed a defect in protein processing that may or may not be responsible for the decreased accumulation of viral RNA observed in constant-temperature infections. 13 ts mutations often affect the folding of mutant proteins (2), B which can result in increased susceptibility of mutant proteins to intracellular proteases. Comparison of the intensities of the viral protein bands in Fig. 5a, labeled at the permissive temperature and subsequently incubated at the permissive and nonpermissive temperatures, reveals no significant change in the half-life of mutant 3D-containing polypeptides at the nonpermissive temperature. ts mutants 3D-106, 3D-107, 3D-108, 3D-109, 3D-110, 3D- 111, 3D-112, and 3D-114 thus display no defects in translation, protein processing, or protein stability that can explain their ts phenotypes. Each of these mutants was studied further to determine whether it displayed a primary defect in RNA synthesis as assayed by temperature shift. 3D-1 13, on the other hand, displayed a defect in protein processing, accumulating a novel polypeptide at both the permissive and nonpermissive I..". - temperatures. Therefore, it would be difficult to establish a primary defect in RNA synthesis for this mutant, and 3D-113 was not studied further. Temperature shift experiments to monitor accumulation of FIG. 4. Effect of temperature shift on accumulaition of newly positive-strand RNA. To determine whether any of the ts synthesized proteins in wild-type and ts mutant-inifected cells at mutants has a primary defect in RNA replication, we again permissive and nonpermissive temperatures. The amou1nt ot tricnioro- labeling cells acetic acid-precipitable radioactivity was determined by infected with wild-type or mutant viruses 7 h postinfection at 32.5 C with [35S]methionine for 15 min at either 32.5 C (lane-s A) or 39.5 C (lanes B) as described in Materials and Methods. The viiral proteins are labeled capsid proteins electrophoresed on the same gels and by identified at left; they were identified by comparing thie mobilities of comparison with similar gels from other laboratories (6, 19). Labeled the labeled viral proteins with those of unlabeled protein markers and proteins from mock-infected cells at 32.5 and 39.5 C are also displayed.

10 872 DIAMOND AND KIRKEGAARD J. VIROL. a Mck Wdtype 3D-107 A B A B s--i Mmc - m _ WdtYP3D-p10 A * A B b A B C D E F a VW ieīl 97.4 kos XCD 3D VP3 5.5 kdad.d 4 VP2 =. VP3 _ EEi. FIG. 5. (a) Effect of temperature shift on protein processing and stability in wild-type and mutant infections. Cells were labeled with [35S]methionine for 15 min at 6 h postinfection at 32.5 C and chased either for 90 min at 32.5 C (lanes A) or for 60 min at 39.5 C (lanes B) as described in Materials and Methods. Labeled proteins from mock-infected cells at 32.5 C are also displayed. (b) SDS-PAGE mobilities of wild-type and selected 3D mutant proteins expressed in E. coli. Mutant and wild-type 3D proteins were expressed from a plasmid, pt5t-3d, constructed by T. Jarvis (Ribozyme Products, Inc.) and similar to a 3D-expression plasmid described previously (52). Plasmid-encoded proteins were selectively labeled with [35S]methionine as described in Materials and Methods. Labeled proteins from cultures of E. coli with no plasmid in the absence (A) and presence (B) of rifampin and from rifampin-treated E. coli containing plasmids encoding wild-type 3D (C and G), 3D protein bearing the 3D-106 mutations (D), 3D protein bearing the 3D-110 mutations (E), and 3D protein bearing the 3D-111 mutations (F) are shown. performed temperature shift experiments in which viral proteins and RNA were allowed to accumulate at the permissive temperature. Then, at 4 h postinfection at 32.5 C, incubations of duplicate cultures were continued at either 32.5 or 39.5 C, samples were taken at intervals, and cytoplasmic extracts were prepared for analysis of RNA accumulation by quantitative dot blot hybridization. The temperature was shifted at an earlier time in the infectious cycle than in the previous temperature shift experiments because of the decreased sensitivity of assays to detect the new synthesis of RNA compared with assays to detect the new synthesis of viral polypeptides. Figure 6a shows that in infections with wild-type poliovirus, more RNA accumulated when the temperature was shifted to 39.5 C than when the incubation was continued at 32.5 C. In infections with ts mutant 3D-107 (Fig. 6b), however, less RNA accumulated after the shift to 39.5 C than when the incubations were continued at the permissive temperature. These data for the 6-h time points of wild-type and 3D-107 infections, as well as for temperature-shifted infections with 3D-106, 3D-109, 3D- 110, 3D-111, 3D-112, 3D-113, and 3D-114, are shown in Fig. 6c. Each mutant, with the exception of 3D-109, showed a significant decrease in RNA accumulation at the nonpermissive condition relative to that of wild-type poliovirus. DISCUSSION Using a clustered charged-to-alanine mutagenesis strategy, in which clusters of codons for charged amino acids are replaced by alanine codons, an array of ts alleles of the 3D coding region was generated. Thirty-seven percent of the mutant cdnas constructed gave rise to mutant viruses with ts phenotypes. However, one of these cdnas contained an additional single-nucleotide substitution (Tables 1 and 2), thus reducing the yield of ts mutants whose phenotypes are known to be caused by the clustered charged-to-alanine mutations to 35%. These results are similar to those achieved with the yeast actin system, in which 44% of the mutations constructed, when present in a single copy in haploid yeast cells, conferred ts phenotypes (76). One difference between these two mutageneses, however, is the amount of lethality observed. Clustered charged-to-alanine mutagenesis of yeast actin gave rise to 32% nonviable haploid yeast cells; our mutagenesis of 3D led to lethal or nearly lethal phenotypes for 59% of the mutations constructed. We do not know whether this results from the RNA-dependent RNA polymerase's being more sensitive to mutation or from the necessity of the viral positive strand to function in capacities other than as the mrna for the mutagenized protein. In any case, clustered charged-to-alanine mutagenesis gave rise to a high percentage of ts mutants, both in genes encoding a structural protein (76) and a protein with enzymatic function. The mutations responsible for the phenotypes of most of these ts mutants lie in the N-terminal third of the 3D coding region, where no well-characterized mutations responsible for viable mutants have been previously identified (Fig. 7). The locations in the 3D coding sequence in which mutations have given rise to viable mutant viruses in other laboratories (1, 10, 18, 19), as well as regions where the 3D sequence exhibits sequence similarity with those of other RNA-dependent RNA polymerases (16, 38, 53), are also shown in Fig. 7. Constant-temperature, single-cycle infections of nine stable ts mutants displayed defects in viral yield or RNA accumulation for eight of the mutants, with the severity of defects mirrored by the severity of plaque defects (Tables 1 and 2). For example, mutant 3D-108, which displayed no defects in viral yield or in RNA accumulation in single-cycle infections, also displayed the least severe small-plaque phenotype, whereas the plaque reduction mutants 3D-107, 3D-112, and 3D-113 all displayed severe defects in both viral yield and RNA accumulation.

11 200 VOL. 68, 1994 CHARGED-TO-ALANINE MUTAGENESIS OF POLIOVIRUS 3D 873 stability compared with wild-type virus. One of the ts mutant 2 > viruses, 3D-113, displayed a defect in the processing of viral 0,' polypeptides. The remaining eight ts mutant viruses, however, z 10000_- showed no defects in viral protein processing. cr,'it was during the analysis of positive-strand RNA accumu- N@,/' lation that most of the ts mutant viruses displayed defects dependent on a temperature shift. Mutants 3D-106, 3D-107, 3D-108, 3D-110, 3D-111, 3D-112, and 3D-114 all exhibited a O '1 / decrease in RNA accumulation compared with that in wild- = -, /type virus 2 h after a shift to the nonpermissive temperature. O 3D-113 was not tested in these experiments because its defect E 1 + > in protein processing would make it difficult to interpret any,-' temperature-dependent effect on RNA accumulation. Of the 0,,, viruses tested, only 3D-109 showed no temperature-dependent defect in RNA accumulation. We do not know whether the < T / simplicity, the mutants resulting from clustered charged-to- Z alanine scanning mutagenesis have been discussed as though 'a the relevant changes were in the amino acid sequence. How-.N ever, ts mutants of poliovirus with defects in RNA accumula- J,T/ tion have been described previously whose only mutations C 1000 /. poliovirus mutants bearing mutations in the coding regions of 0 E / 2B (46), 3A (27), and 3D (10) owe their mutant phenotypes to with the S. cerevisiae ACT] mutants (76), it is likely that the Time post infection (hours) 3D-107, 3D-108, 3D-110, 3D-111, 3D-112, and 3D-114, dis- C C.o played ts plaque phenotypes that correlated with ts defects in FIG. 6. Effect of temperature shift on accumulation of positive-strand.5 T :-- RNA in infections with wild-type and ts mutant polioviruses. Means iu -and standard deviations are shown. (a) HeLa cells were infected with.o300.2x --. wild-type poliovirus at 32.5 C. At 4 h postinfection, the incubations were continued at either 32.5 C (solid line) or 39.5 C (dashed lines),... and the frssrw subsequent... - accumulation of positive-strand RNA was moni- LO sf>.ss v I tored by dot blot hybridization. (b) Accumulation of positive-strand Xn x ;RNA x in T Rinfections S.-.. with ts mutant virus 3D-107 during continuous 100 incubation at 32.5 C (solid line) and following a temperature shift to 0 f'a l l m S>-: l >.5N'S l ss 'SS l sss l 39.5 C (dashed line) at 4 h postinfection. (c) Graphic representation of C the relative accumulation of positive-strand RNA in wild-type and ts 0 mutant-infected cells with and without temperature shift. For each 0 & 0 virus, the accumulation of RNA at 6 h postinfection, following 0 a a0 0 0 continuous incubation at 32.5 C, was normalized to 100% (open bars). The shaded bars show the relative amounts of positive-strand RNA Virus that accumulated by the 6-h time point when the temperature was shifted to 39.5 C at 4 h postinfection. Temperature shift experiments allowed us to identify the a first step in viral replication after cell entry in which each 15000, mutant displayed a biochemical defect. None of the ts mutant viruses displayed any defects in translation or in protein failure to display a defect in RNA accumulation after the temperature shift resulted from the subtlety of the mutant phenotype or from a defect elsewhere in the viral replicative b 4000 cycle, for example, in establishing but not maintaining the replication complex (48). Each set of mutations introduced created changes in both the positive- and negative-strand viral RNAs as well as in the amino acid sequences of 3D and related polypeptides. For ->_-f-- > s were in the 3' noncoding region (61) or the 5' noncoding o 1, region (3, 24, 57). Furthermore, it is possible that cis-dominant < l g defects in RNA structure or sequence. Experiments to test the recessive, cis-dominant, or dominant nature of these mutant 0 phenotypes have not been performed; however, by analogy defects caused by each of these sets of mutations are due, at least in part, to the mutations in the protein sequence. Seven of the ts mutants generated in this study, 3D-106, t5.rna accumulation after temperature shift. The mutations c_ responsible for the ts phenotype of 3D-114 lie near the C

12 874 DIAMOND AND KIRKEGAARD J. VIROL. 3D D-106 3D-108 K>i 3D-110 3D-111 N L 11 L 1 11 I -K -I 1 I -1 -i -WIII -1 IF I I I D-112 lii I I-I II III II I I J.- 1,, - i t 250 I 1 I Se1-D-18 III 1' IF n -II i 3D I- 1 -l I 1 L 1 I 1 I 1 vehl-3d-7256(n-h) 3D-3 vehl-3d-7256(n-d) vehl -3D-7256(N-Y) FIG. 7. Schematic representation of the 461-amino-acid sequence of 3D, the poliovirus RNA-dependent RNA polymerase, showing the locations of charged residues and clustered charged-to-alanine mutations. The locations of acidic residues are shown by short vertical slashes below the horizontal line that represents the 461-amino-acid sequence; basic residues are shown by slashes above the line. The locations of the mutations in 3D-106, 3D-107, 3D-108, 3D-110, 3D-111, 3D-112, and 3D-114, those ts mutants that displayed specific defects in RNA accumulation in temperature shift experiments, are shown as follows: 0, ts virus with a plaque reduction phenotype; O, ts virus with a small-plaque phenotype. Regions of homology with other RNA-dependent RNA polymerases (16, 38, 53) are depicted by brackets. Region A, amino acids 225 to 240; region B, amino acids 279 to 305; region C: amino acids 323 to 333; region D, amino acids 349 to 361. The locations of mutations that have been previously demonstrated to give rise to viable viruses with conditionally defective mutant phenotypes are indicated by arrows (1, 10, 18, 19). terminus of the 3D protein (Fig. 7), only two amino acids away from Asn-424; mutations in Asn-424 are responsible for the ts phenotypes of three mutants studied previously (1, 18). One of these mutants, veh1-3d-7256(n-h), showed a lower ratio of positive to negative strands in constant-temperature infections performed at high temperature than in those at low temperature, suggesting a specific defect in positive-strand synthesis at the nonpermissive temperature (18). It will be interesting to test whether 3D-114 displays a specific defect in positive-strand synthesis in temperature shift experiments. The mutations responsible for the ts phenotypes of 3D-106, 3D-107, 3D-108, 3D-110, 3D-111, and 3D-112 all lie in the N-terminal third of the 3D coding region (Fig. 7). No viable mutants with mutations in this region of 3D have been identified previously, and this region displays no sequence similarity with other RNAdependent RNA polymerases (16, 38, 53). The most severe biochemical and plaque morphology phenotypes were exhibited by ts mutants 3D-107, 3D-112, and 3D These mutants are quite suitable for genetic studies that require pronounced plaque phenotypes. For yeast actin, it was shown that clustered charged-to-alanine mutagenesis specifically targeted patches of charged residues on the protein surface (76). For tissue plasminogen activator, 90% of the mutant proteins that resulted from a similar mutagenesis protocol were stable and properly folded when expressed in bacteria (8). As discussed in the introduction, it is possible that the reason this mutagenesis algorithm can generate ts mutants at such a high frequency in actin, and in the 3D polymerase, is that the targeted clusters of surface charges are normally involved in hydrogen bonding or electrostatic interactions with water or other proteins (2) and that these interactions are crucial for protein folding, function, or both. If this is the case, suppression analysis of mutant alleles generated by clustered charged-to-alanine mutagenesis may prove especially fruitful to identify proteins that interact functionally with the wild-type proteins (76). In the case of 3D, suppression analysis (3, 4, 24, 34, 47) of these alleles may identify other components of the poliovirus replication complex that interact functionally with the RNA-dependent RNA polymerase during the poliovirus infectious cycle. ACKNOWLEDGMENTS We thank Peter Sarnow, Mary T. L. Beckman, and Janice A. Pata for helpful comments on the manuscript and David Botstein for communication of results prior to publication. S.E.D. thanks Ben Young for advice on constructing mutant cdnas. This research was supported by NIH grant AI and by the David and Lucile Packard Foundation. S.E.D. was supported by fellowships from the NSF and the Colorado Institute for Research in Biotechnology. We are grateful to the Keck Foundation for generous support of RNA research in Boulder. K.K. is an assistant investigator of the Howard Hughes Medical Institute. REFERENCES 1. Agut, H., K. M. Kean, 0. Fichot, J. Morasco, J. B. Flanegan, and M. Girard A point mutation in the poliovirus polymerase gene determines a complementable temperature-sensitive defect of RNA replication. Virology 168: Alber, T Mutational effects on protein stability. Annu. Rev. Biochem. 58: Andino, R., G. E. Rieckhof, and D. Baltimore A functional ribonucleoprotein complex forms around the 5' end of poliovirus RNA. Cell 63: Andino, R., G. E. Rieckhof, D. Trono, and D. Baltimore Substitutions in the protease 3C gene of poliovirus can suppress a mutation in the 5' noncoding region. J. Virol. 64: Argos, P A sequence motif in many polymerases. Nucleic Acids Res. 16: Barton, D. J., and J. B. Flanegan Coupled translation and replication of poliovirus RNA in vitro: synthesis of functional 3D polymerase and infectious virus. J. Virol. 67: Bass, S. H., M. G. Mulkerrin, and J. A. Wells A systematic mutational analysis of hormone-binding determinants in the human growth hormone receptor. Proc. NatI. Acad. Sci. USA 88: Bennett, W. F., N. F. Paoni, B. A. Keyt, D. Botstein, J. J. S. Jones, L. Presta, F. M. Wurm, and M. J. Zoller High resolution analysis of functional determinants on human tissue-type plasminogen activator. J. Biol. Chem. 266:

13 VOL. 68? 1994 CHARGED-TO-ALANINE MUTAGENESIS OF POLIOVIRUS 3D Bernstein, H. D., and D. Baltimore Poliovirus mutant that contains a cold-sensitive defect in viral RNA synthesis. J. Virol. 62: Bernstein, H. D., P. Sarnow, and D. Baltimore Genetic complementation among poliovirus mutants derived from an infectious cdna clone. J. Virol. 60: Bienz, K., D. Egger, T. P(jister, and M. Troxler Structural and functional characterization of the poliovirus replication complex. J. Virol. 66: Bienz, K., D. Egger, Y. Rasser, and W. Bossart Kinetics and location of poliovirus macromolecular synthesis in correlation to virus-induced cytopathology. Virology 100: Bienz, K., D. Egger, M. Troxler, and L. Pasamontes Structural organization of poliovirus RNA replication is mediated by viral proteins of the P2 genomic region. J. Virol. 64: Blair, W. S., and B. L. Semler Role for the P4 amino acid residue in substrate utilization by the poliovirus 3CD proteinase. J. Virol. 65: Blumenthal, T., T. A. Landers, and K. Wever Bacteriophage Q1 replicase contains the protein biosynthesis elongation factors EF Tu and EF Ts. Proc. Natl. Acad. Sci. USA 69: Bruenn, J. A Relationships among the positive strand and double-strand RNA viruses as viewed through their RNA-dependent RNA polymerases. Nucleic Acids Res. 19: Burns, C. C., M. A. Lawson, B. L. Semler, and E. Ehrenfeld Effects of mutations in poliovirus 3Dpol on RNA polymerase activity and on polyprotein cleavage. J. Virol. 63: Burns, C. C., 0. C. Richards, and E. Ehrenfeld Temperature-sensitive polioviruses containing mutations in RNA polymerase. Virology 189: Charini, W. A., C. C. Burns, E. Ehrenfeld, and B. L. Semler trans rescue of a mutant poliovirus RNA polymerase function. J. Virol. 65: Cunningham, B. C., and J. A. Wells High-resolution epitope mapping of hgh-receptor interactions by alanine-scanning mutagenesis. Science 244: Dao-pin, S., D. E. Anderson, W. A. Baase, F. W. Dahlquist, and B. W. Matthews Structural and thermodynamic consequences of burying a charged residue within the hydrophobic core of T4 lysozyme. Biochemistry 30: Dasgupta, A., M. H. Baron, and D. Baltimore Poliovirus replicase: a soluble enzyme able to initiate copying of poliovirus RNA. Proc. Natl. Acad. Sci. USA 76: Delarue, M., 0. Poch, N. Tordo, D. Moras, and P. Argos An attempt to unify the structure of polymerases. Protein Eng. 3: Dildine, S. L., and B. L. Semler The deletion of 41 proximal nucleotides reverts a poliovirus mutant containing a temperaturesensitive lesion in the 5' noncoding region of genomic RNA. J. Virol. 63: Dorssers, L., S. van der Krol, J. van der Meer, A. van Kammen, and P. Zabel Purification of cowpea mosaic virus RNA replication complex: identification of a virus-encoded 110,000- dalton polypeptide responsible for RNA chain elongation. Proc. Natl. Acad. Sci. USA 81: Fujimura, T., and R. B. Wickner Reconstitution of template-dependent in vitro transcriptase activity of a yeast doublestranded RNA virus. J. Biol. Chem. 264: Giachetti, C., S. S. Hwang, and B. L. Semler cis-acting lesions targeted to the hydrophobic domain of a poliovirus membrane protein involved in RNA replication. J. Virol. 66: Giachetti, C., and B. L. Semler Role of a viral membrane polypeptide in strand-specific initiation of poliovirus RNA synthesis. J. Virol. 65: Gibbs, C. S., and M. J. Zoller Rational scanning mutagenesis of a protein kinase identifies functional regions involved in catalysis and substrate interactions. J. Biol. Chem. 266: Grakoui, A., R. Levis, R. Raju, H. V. Huang, and C. Rice A cis-acting mutation in the Sindbis virus junction region which affects subgenomic RNA synthesis. J. Virol. 63: Hayes, R. J., and K. W. Buck Complete replication of a eukaryotic virus RNA in vitro by a purified RNA-dependent RNA polymerase. Cell 63: Jablonski, S. A., M. Luo, and C. D. Morrow Enzymatic activity of poliovirus RNA polymerase mutants with single amino acid changes in the conserved YGDD amino acid motif. J. Virol. 65: Jablonski, S. A., and C. D. Morrow Enzymatic activity of poliovirus RNA polymerases with mutations at the tyrosine residue of the conserved YGDD motif: isolation and characterization of polioviruses containing RNA polymerases with FGDD and MGDD sequences. J. Virol. 67: Jacobson, S. J., D. A. M. Konings, and P. Sarnow Biochemical and genetic evidence for a pseudoknot structure at the 3' terminus of the poliovirus RNA genome and its role in viral RNA amplification. J. Virol. 67: Jarvis, T., and K. Kirkegaard Poliovirus RNA recombination: mechanistic studies in the absence of selection. EMBO J. 11: Johnson, K., and P. Sarnow Three poliovirus 2B mutants exhibit noncomplementable defects in viral RNA amplification and display dosage-dependent dominance over wild-type poliovirus. J. Virol. 65: Jore, J., B. De Gues, R. J. Jackson, P. H. Pouwels, and B. E. Enger-Valk Poliovirus protein 3CD is the active protease for processing of the precursor protein in vitro. J. Gen. Virol. 69: Kamer, G., and P. Argos Primary structural comparison of RNA-dependent polymerases from plant, animal and bacterial viruses. Nucleic Acids. Res. 12: Kanaya, S., N. C. Katsuda, and M. Ikehara Importance of the positive charge cluster in Escherichia coli ribonuclease HI for the effective binding of the substrate. J. Biol. Chem. 266: Kirkegaard, K., and D. Baltimore The mechanism of RNA recombination in poliovirus. Cell 47: Kirkegaard, K., and B. Nelsen Conditional poliovirus mutants made by random deletion mutagenesis of infectious cdna. J. Virol. 64: Kitamura, N., B. Semler, P. Rothberg, G. Larsen, C. Adler, A. Dorner, E. Emini, R. Hanecak, J. Lee, S. van der Werf, C. Anderson, and E. Wimmer Primary structure, gene organisation, and polypeptide expression of poliovirus RNA. Nature (London) 291: Kunkel, T. A Rapid and efficient site-specific mutagenesis without phenotypic selection. Proc. Natl. Acad. Sci. USA 82: Laemmli, U. K Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature (London) 227: La Monica, N., C. Meriam, and V. R. Racaniello Mapping of sequences required for mouse neurovirulence of poliovirus type 2 Lansing. J. Virol. 57: Li, J.-P., and D. Baltimore Isolation of poliovirus 2C mutants defective in viral RNA synthesis. J. Virol. 62: Li, J.-P., and D. Baltimore An intragenic revertant of a poliovirus 2C mutant has an uncoating defect. J. Virol. 64: Maynell, L. A., K. Kirkegaard, and M. W. Klymkowsky Inhibition of poliovirus RNA synthesis by brefeldin A. J. Virol. 66: Mills, D. R., C. Priano, P. DiMauro, and B. D. Binderow Q0 replicase: mapping the functional domains of an RNA-dependent RNA polymerase. J. Mol. Biol. 205: a.Molla, A., A. Paul, and E. Wimmer. Personal communication. 50. Molla, A., A. V. Paul, and E. Wimmer Cell-free, de novo synthesis of poliovirus. Science 254: Morrow, C. D., B. Warren, and M. R. Lentz Expression of enzymatically active poliovirus RNA-dependent RNA polymerase in Escherichia coli. Proc. Natl. Acad. Sci. USA 84: Plotch, S. J., 0. Palant, and Y. Gluzman Purification and properties of poliovirus RNA polymerase expressed in Eschericlhia coli. J. Virol. 63: Poch, O., I. Sauvaget, M. Delarue, and N. Tordo Identification of four conserved motifs among the RNA-dependent polymerase encoding elements. EMBO J. 8:

14 876 DIAMOND AND KIRKEGAARD 54. Quadt, R., C. Koa, K. Borwinging, R. Hershberger, and P. Ahlquist Characterization of a host protein associated with brome mosaic virus RNA-dependent RNA polymerase. Proc. Natl. Acad. Sci. USA 90: Racaniello, V. R., and D. Baltimore Cloned poliovirus complementary DNA is infectious in mammalian cells. Science 214: Racaniello, V. R., and D. Baltimore Molecular cloning of poliovirus cdna and determination of the complete nucleotide sequence of the viral genome. Proc. Natl. Acad. Sci. USA 78: Racaniello, V. R., and C. Meriam Poliovirus temperaturesensitive mutant containing a single nucleotide deletion in the 5' noncoding region of the viral RNA. Virology 155: Ribas, J. C., and R. B. Wickner RNA-dependent RNA polymerase consensus sequence of the L-A double-stranded RNA virus: definition of essential domains. Proc. Natl. Acad. Sci. USA 89: Sankar, S., and A. G. Porter Point mutations which drastically affect the polymerization activity of encephalomyocarditis virus RNA-dependent RNA polymerase correspond to the active site of Escherichia coli DNA polymerase. J. Biol. Chem. 267: Sarnow, P Role of 3'-end sequences in infectivity of poliovirus transcripts made in vitro. J. Virol. 63: Sarnow, P., H. D. Bernstein, and D. Baltimore A poliovirus temperature-sensitive RNA synthesis mutant located in a noncoding region of the genome. Proc. Natl. Acad. Sci. USA 83: Semler, B. L., R. J. Kuhn, and E. Wimmer Replication of the poliovirus genome, p In E. Domingo, J. J. Holland, and P. A. Ahlquist (ed.), RNA genetics, vol. 1. CRC Press, Boca Raton, Fla. 63. Sonenberg, N Regulation of translation by poliovirus. Adv. Virus Res. 33: Steinhauer, D. A., and J. J. Holland Rapid evolution of RNA viruses. Annu. Rev. Microbiol. 41: Strauss, J. H., and E. G. Strauss Evolution of RNA viruses. Annu. Rev. Microbiol. 42: Studier, F. W., and B. A. Moffat Use of bacteriophage T7 RNA polymerase to direct selective high-level expression of cloned genes. J. Mol. Biol. 189: J. VIROL. 67. Studier, F. W., A. H. Rosenberg, J. J. Dunn, and J. W. Dubendorff Use of T7 RNA polymerase to direct the expression of cloned genes. Methods Enzymol. 185: Thukral, S. K., M. L. Morrison, and E. T. Young Alanine scanning site-directed mutagenesis of the zinc fingers of transcription factor ADR1: residues that contact DNA and that transactivate. Proc. Natl. Acad. Sci. USA 88: Traynor, P., B. M. Young, and P. Ahlquist Brome mosaic virus 2a protein: effects on RNA replication and systematic spread. J. Virol. 65: Trono, D., R. Andino, and D. Baltimore An RNA sequence of hundreds of nucleotides at the 5' end of poliovirus RNA is involved in allowing viral protein synthesis. J. Virol. 62: Tuschall, D. M., E. Hiebert, and J. B. Flanegan Poliovirus RNA-dependent RNA polymerase synthesizes full-length copies of poliovirion RNA, cellular mrna, and several plant virus RNAs in vitro. J. Virol. 44: van der Werf, S., J. Bradley, E. Wimmer, F. W. Studier, and J. J. Dunn Synthesis of infectious poliovirus RNA by purified T7 RNA polymerase. Proc. Natl. Acad. Sci. USA 83: Van Dyke, T. A., and J. B. Flanegan Identification of poliovirus polymerase P63 as a soluble RNA-dependent RNA polymerase. J. Virol. 35: Wahba, A. J., M. J. Miller, A. Niveleau, T. A. Landers, G. G. Carmichael, K. Weber, D. A. Hawley, and L. I. Slobin Subunit I of Q,B replicase and 30S ribosomal protein S1 of Escherichia coli. Evidence for the identity of the two proteins. J. Biol. Chem. 249: Ward, C. D., M. A. Stokes, and J. B. Flanegan Direct measurement of the poliovirus RNA polymerase error frequency in vitro. J. Virol. 62: Wertman, K. F., D. G. Drubin, and D. Botstein Systematic mutational analysis of the yeast ACT1 gene. Genetics 132: Wu, S.-X., P. Ahlquist, and P. Kaesberg Active complete in vitro replication of nodavirus RNA requires glycerophospholipid. Proc. Natl. Acad. Sci. USA 89: Ypma-Wong, M. F., P. G. Dewalt, V. H. Johnson, J. G. Lamb, and B. L. Semler Protein 3CD is the major poliovirus proteinase responsible for cleavage of the P1 capsid precursor. Virology 166: